Lionfish Dissection: Techniques and Applications
NOAA Technical Memorandum NOS NCCOS 139
Mention of trade names or commercial products does not constitute
endorsement or recommendation for their use by the United States
government.
Citation for this Report
Green, S.J., Akins, J.L., and J.A. Morris, Jr. 2012. Lionfish dissection: Techniques and
applications. NOAA Technical Memorandum NOS NCCOS 139, 24 pp.
Lionfish Dissection: Techniques and Applications
Stephanie J. Green
Department of Biological Sciences
Simon Fraser University
8888 University Drive
Burnaby, British Columbia
Canada, V5A 1S6
John L. Akins
Reef Environmental Education Foundation
98300 Overseas Highway
Key Largo, Florida 33037
James A. Morris, Jr., Ph.D.
Center for Coastal Fisheries and Habitat Research
NOAA/NOS/NCCOS
101 Pivers Island Road
Beaufort, North Carolina 28516
NOAA Technical Memorandum NOS NCCOS 139
2012
United States Department of
Commerce
National Oceanic and
Atmospheric Administration
National Ocean Service
John E. Bryson
Acting Secretary
Jane Lubchenco
Administrator
David M. Kennedy
Assistant Administrator
Table of Contents
INTRODUCTION ...........................................................................................................................1
EUTHANASIA AND ANIMAL WELFARE .................................................................................2
HANDLING PROCEDURES AND PRECAUTIONS ...................................................................3
Dissection.............................................................................................................................3
Specimen disposal ................................................................................................................3
First aid for envenomation ...................................................................................................3
MEASUREMENT AND SAMPLING OVERVIEW......................................................................4
Minimizing measurement error............................................................................................4
External anatomy .................................................................................................................6
Internal anatomy ..................................................................................................................7
DISSECTION METHODOLOGY ..................................................................................................8
External measurements and sampling..................................................................................8
Weight ......................................................................................................................8
Length ......................................................................................................................8
Gape .........................................................................................................................9
Tissue sampling .......................................................................................................9
Internal measurements and sampling .................................................................................10
Opening the gut cavity ...........................................................................................10
Gender identification and reproductive staging .....................................................12
Interstitial fat deposits ............................................................................................14
Stomach contents ...................................................................................................14
Otolith removal ......................................................................................................16
ACKNOWLEDGEMENTS ...........................................................................................................19
LITERATURE CITED ..................................................................................................................20
APPENDICES ...............................................................................................................................21
I. Lionfish dissection specimen data sheet template .......................................................21
II. Lionfish stomach contents data sheet template ............................................................22
III. Dissection equipment list .............................................................................................23
IV. Lionfish tissue repository.............................................................................................24
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List of Figures
Fig. 1. Lionfish external anatomy….………………………………..…………………………….6
Fig. 2. Lionfish internal anatomy…....………………………….…………………………………7
Fig. 3. Taking external lionfish measurements: A) Lionfish weight measurement. Ensure
the specimen is thoroughly blotted dry and that the entire specimen is balanced on the
scale. B) Measuring lionfish total and standard length to the nearest 1 mm. The arrow
indicates the location of the base of the caudal fin...……………..……………………….………9
Fig. 4. Measuring lionfish A) gape height and B) gape width to the nearest 1 mm. Ensure
that the specimen’s mouth is open to its fullest extent..…………………………………………..9
Fig. 5. Sample collection of three tissue types for genetic and stable isotope research: A) gill,
B) pectoral fin, and C) muscle tissue. Sample size for gill and fin tissue is a 1-cm2
section; muscle tissue sample is a 1-cm3 section...………………………………....………...…10
Fig. 6. Opening the lionfish’s gut cavity. A) A shallow incision extends from the
urogenital opening towards the base of the pelvic fins (pelvic girdle). B) As you approach
the pelvic fins, a deeper cut will be required to sever the pelvic girdle. C) Cutting along the
rear edge of the gill arch toward the dorsal fin. D) Lifting the flank to expose the gut cavity
and internal organs, cutting through any minor connecting tissue as needed.………………..….11
Fig. 7. A) Removal of interstitial fat deposits from intestines and B) measuring interstitial
fat volume to nearest 1 ml...…………………………...…………....……………………………14
Fig. 8. Stomach dissection and contents analysis. A) Using dissecting scissors sever the
esophagus where it terminates. B) Insert scissors into the esophageal opening of the stomach.
C) Make a shallow cut along the length of the stomach wall to the intestine opening.
D) Invert the stomach and remove all prey items; placing prey onto a clean surface
for sorting and identification…………….....……………….……………………………………15
Fig. 9. Ontogenetic change in sagittal otolith shape, taken from lionfish of A) 250 mm,
B) 325 mm and C) 357 mm total length.………………………….…...………………………...16
ii
Fig. 10. Sagittal otolith removal. A) Severe the spinal cord between the head and the
first dorsal fin ray. B) Grasp the gill filaments and arches and severe their attachment to
the roof of the oral cavity. C) Locate the cranial cavity by running two fingers along both
sides of the spinal cord anteriorly until a slight bulge is felt. D) Make a cut on the posterior
side of the cavity down through the spine. This incision allows access to the otic bulba,
which contains the sagittal otoliths. E) Brain matter (white, soft tissue) will often be
exposed by opening the cranial cavity. F) Reach gently into the cranial cavity to remove
the free-floating otoliths…………………………………….……………………………………18
List of Tables
Table 1. Dissection activities and some of the primary uses of data generated by each type
of measurement or sample. Scientific measurements are generally expressed in metric units,
allowing for universal comparison (i.e., millimeters [mm], milliliters [ml] and grams [g])...…..4
Table 2. Sex determination and gonad staging key for lionfish..………………………….…….13
Table 3. Five-point scale for scoring the digestion of prey items removed from a lionfish
stomach.……………………………………………………………………………………….…16
iii
INTRODUCTION
The Goals of Dissection
Dissection can provide unique information on the physiology, biology and ecology of
organisms. This document describes protocols for dissecting lionfish (Pterois volitans and P.
miles). Protocols were developed to provide guidance to trained research personnel. Lionfish
are native to the Indo-Pacific, but have become established in marine habitats within the Western
Atlantic, Gulf of Mexico and Caribbean. The protocols described within this document were
designed to help standardize handling and dissection methodologies for these species, with the
goal of improving the coordination of research (e.g., Lionfish Tissue Repository; Appendix V).
We focus on dissection methods, which yield data that contribute to our understanding of
lionfish biology and ecology. By pairing dissection information with environmental and biotic
data, researchers and managers can better understand lionfish population structure and dynamics,
age and growth, reproductive biology, and food web ecology on various temporal and spatial
scales.
Population structure
The structure of populations can be determined through genetic approaches, which use
tissue samples collected from dorsal fin, caudal fin, gill, and muscle tissues.
Population dynamics
Age and growth information from individual fish can be determined using otoliths (ear
stones) in conjunction with fish body lengths. The relationship between weight and length of
lionfish specimens can also be used to monitor changes in body condition, which may correlate
with environmental variables such as: habitat type, resource availability, changing latitude, or
invasion status. These data can also be used to determine temporal and spatial changes in age
structure of local populations.
Reproductive biology
The reproductive status of an individual lionfish can be determined using various metrics
such as gonad weight, morphology, and histology. These data aggregated over space and time
can yield population-level trends in reproductive biology, such as spawning frequency and
seasonality, and maturation schedules.
Food web ecology
Lionfish diet can be characterized by identifying and measuring the stomach contents of
collected specimens. Conducting stomach contents analysis across temporal and spatial scales
can improve our understanding of changes in diet composition and potential community effects.
The carbon and nitrogen isotopic signature of lionfish muscle and fin tissues can be used to
assess trophic position and feeding ecology. These data can be used in conjunction with isotope
1
data from native species to describe biotic interactions between lionfish and members of the
invaded food web.
EUTHANASIA AND ANIMAL WELFARE
Many organizations such as universities, government agencies and scientific journals have
strict standards for the humane treatment of animal taxa used in scientific study. These standards
apply to methods by which death is caused in specimens destined for dissection. Euthanasia is
the act of inducing humane death in an animal (American Veterinary Medical Association,
2007). At all times, research personnel should ensure that euthanasia is performed in an ethical
manner to minimize pain and distress to the specimen. Euthanasia of lionfish specimens can be
achieved through several methods, depending on the resources available and the intended use of
the fish.
Euthanasia methods
The following euthanasia methods are appropriate for live-captured specimens. For
information on techniques for live capture please refer to Akins (in press).
Eugenol
If lionfish specimens are not destined for human consumption, they can be euthanized
using an overdose of eugenol (clove oil and isopropyl alcohol; in accordance with Borski and
Hodson 2003). Typical response includes rapid swimming followed by loss of equilibrium and
inverted floating at the surface. Death is marked by the cessation of opercular (gill) movement
for a period exceeding 5 minutes. Clove oil overdose refers to the exposure of a fish to a
concentration greater than that required for anaesthesia ultimately resulting in death. These doses
have been described in the literature for numerous body sizes and fish species (Borski and
Hodson 2003; Neiffer et al. 2004; Soltani et al. 2004). Collectively or individually, fish should
be transferred directly to a solution of 10 L seawater and 50 ml/L eugenol (a mixture of 1/10
clove oil and 9/10 alcohol by volume; 5 ml clove oil and 45 ml alcohol per liter of seawater or 50
ml clove oil and 450ml for 10L seawater). Lionfish response to this treatment is rapid; cessation
of gill movement generally occurs within 5 minutes of exposure to the solution.
Ice slurry
In cases where the lionfish meat is destined for human consumption, the handler may
immerse the lionfish in an ice-seawater slurry for approximately 15 min. Exposure to low
temperatures decreases the heart rate of lionfish to the point of death (Wilson et al. 2009;
Blessing et al. 2010).
2
HANDLING PROCEDURES AND PRECAUTIONS
During dissection
Lionfish spines can remain venomous for some time after death (depending on
temperature) and those who handle lionfish must take extra precautions to protect themselves
from envenomation (Badillo et al. 2012). Aside from their venomous nature, a lionfish’s dorsal,
pelvic and anal spines can pose a risk because they are needle sharp (Fig. 1). When handling
both live and euthanized lionfish, it is prudent to wear puncture resistant gloves to reduce the risk
of envenomation. The symptoms of envenomation include pain and swelling in the affected
appendage, but could include sweating, respiratory distress and even temporary paralysis in more
severe cases (Badillo et al. 2012). Puncture resistant gloves manufactured by Hex Armor®,
TurtleSkin®, and other specialty companies are designed specifically to prevent needle-stick and
other types of industrial punctures. Heavy leather or canvas work gloves, latex or nitrile
examination gloves are not suitable substitutes as lionfish spines can easily penetrate these
materials. When using protective equipment, follow the manufacturer’s instructions and be
cognizant of the area of protection. Some gloves may only provide protection in the palm and
fingers.
Before dissection begins (following completion of any weight and length measurements),
the lionfish’s spines can be removed with a sharp pair of scissors or knife. Cutting the spines off
at their base can minimize risk of envenomation. Protection against skin irritation caused by the
non-venomous spines (opercular spines or cheek mail; Fig. 1H) on lionfish heads can be
provided by wearing protective gloves made of nitrile, latex, or more durable rubber compounds.
Specimen disposal
Following dissection, care should be taken to ensure that lionfish carcasses, including
spines, are safely handled and disposed. Discarded lionfish parts that contain venomous spines
should be placed in stout, puncture-proof, and capped containers such as a glass or plastic jar or
bottle to avoid possible envenomation during waste handling. Carcasses containing venomous
spines should not be placed back into near-shore or high visitation marine environments as
currents may bring the carcasses back into public areas.
First aid for envenomation
In the event of a lionfish sting, immediate first aid treatment includes immersion of the
affected area in non-scalding hot water until pain subsides. Secondary medical treatment may be
advised, as symptoms and complications vary depending on the severity of the sting, whether or
not part of the spine remains in the wound, and individual reactions to lionfish venom. Further
information on lionfish stings and their treatment can be obtained at http://www.reef.org/lionfish
(REEF 2012) or http://www.ccfhr.noaa.gov/stressors/lionfsih.aspx (NOAA 2012).
3
MEASUREMENT AND SAMPLING OVERVIEW
This dissection protocol is comprised of a series of measurement and sampling
methodologies, most easily conducted in a specific sequence. We broadly group these
methodologies by ‘external’ and ‘internal’ sampling. A quick reference guide is provided in
Table 1 with additional detail in the following sections.
Table 1. Dissection activities and some of the primary uses of data generated by each type of
measurement or sample from juvenile and adult lionfish. Scientific measurements are generally
expressed in metric units, allowing for universal comparison (i.e., millimeters [mm], milliliters
[ml] and grams [g]).
Type
Metric
Total length (TL)
Measurement Standard length (SL)
Weight
Gape width
Gape height
External
Gill tissue
Units
mm
mm
g
mm
mm
-
Sample
Muscle tissue
-
Fin clip
Gender
Internal
Measurement Interstitial fat volume
Stomach contents
Otoliths
Sample
Gonad
Stomach contents
M or F
ml
mm & ml
-
Application
Growth, body condition,
population size structure
Feeding ecology
Species identification,
population structure
Food web ecology through
stable isotope analysis, species
identification, population
structure
Species identification,
population structure
Individual gender, gender ratio
in population
Health assessment
Feeding ecology
Age and growth
Reproductive physiology
Feeding ecology
4
Minimizing measurement error
Small amounts of measurement error can greatly influence the biological interpretation of
data collected during dissection. It is important that the dissector execute each measurement
carefully, paying close attention to detail, in order to minimize error. When possible, having a
separate data recorder increases the efficiency of the data collection process and reduces
recording error. When a separate recorder is used, verbally confirming each measurement by
repeating it back to the dissector ensures the highest quality data. Original hard copies of data
entry should always be archived for future reference. In all cases, the units of measurement
should be recorded (e.g., mm, g, ml). Pre-labeling all sample vials and bags with sample
identification improves efficiency and decreases cross-contamination. Additional sources of
error, specific to each measurement are described in the following section. Sample data sheets
are provided in Appendices I and II.
5
EXTERNAL ANATOMY
Fig 1. Lionfish external anatomy. Photo credit C. Calloway (main photo).
Code Description
Venomous
Notes
A
Dorsal spines
Yes
13 spines, all are venomous
B
Soft dorsal
No
Secondary dorsal fin; non-venomous
C
Caudal fin
No
Non-venomous
D
Pectoral fin
No
Non-venomous
E
Pelvic fin
Yes
First spine on each pelvic fin is venomous
F
Anal fin
Yes
The first three spines of the anal fin are
venomous
G
Super-ocular
tentacles
No
Missing in this specimen; left inset
H
Opercular spines
No
Larger individuals may have prominent bony
protrusions on the operculum and head;
right inset, also referred to as "cheek mail"
6
INTERNAL ANATOMY
Fig 2. Lionfish internal anatomy.
Code Description
Notes
A
Gill rakers and filaments
B
Swim bladder
Often over inflated in specimens brought up from depth;
may be deflated in speared fish
C
Swim bladder muscles
Used to control position of swim bladder, often confused
with gonads in smaller specimens
D
Liver
Color can be an important indicator of health
E
Gonad
(this specimen: testis)
Often lay along inflated swim bladder
Two gonads; second in same position on the other side of
the body; Males: testes are thin and elongated with edges
Females: ovaries are rounded. Size depends on
development (see additional photographs figures below)
F
Urinary bladder
Clear membrane, usually only visible when filled with
urine; Easily ruptured during dissection
G
Interstitial fatty deposits
White tissue, often surrounding stomach and intestines
H
Stomach
Can expand up to ~30 times when full
I
Intestine
Often contains dark-colored digested matter
7
DISSECTION METHODOLOGY
External measurements and sampling
Weight
Before weighing the lionfish, place your weight boat or specimen tray on the scale and tare
it. Blot the lionfish dry using paper towel, as excess water can be a large source of error. If the
specimen has been euthanized in ice slurry, check the mouth cavity for any remaining ice. When
balancing the lionfish on the scale, ensure that no portion of the fish is touching the table (Fig.
3A). Read and record the unit weight measurement in grams.
Note: When possible, do not take weight measurements on a moving surface (i.e., boat) as the
movement of the vessel will make accurate weight measures difficult.
Length
Use a metric fish board or metric measuring tape secured to a flat surface for all length
measurements, and be sure to place the lionfish ON TOP of the measuring tape. The tape should
run parallel to the fish’s midline from snout to tail (Fig. 3B). Placing the measuring tape on top
of the fish introduces measurement error due to curvature of the tape. To measure total length
(TL), obtain fish length from the tip of the snout to the longest point on the tail (caudal fin),
recording to the nearest 1 mm (Fig. 3B). Ensure that the specimen’s mouth is closed during
measurement. Measure standard length (SL) as the length from the tip of the snout to the last
vertebrae to the nearest 1 mm. The last vertebra is at the base of the end of the fleshy portion of
the caudal peduncle (Fig. 3B) and can be located with manual probing or by flexing the caudal
peduncle to find the last vertebrae.
Note: Accuracy of weight and length measurements may decrease once fish body cavities have
been opened or altered (through filleting or other handling procedures). These conditions should
be noted in any altered specimens.
8
Fig. 3. A) Lionfish weight measurement. Ensure that the specimen is thoroughly blotted dry and
that the entire specimen is balanced on the scale. B) Measuring lionfish total and standard length
to the nearest 1 mm. The arrow indicates the location of the base of the caudal fin.
Mouth gape
To measure gape size, open the mouth of the lionfish to its fullest extent (without
overextending). With a ruler, measure the distance (to the nearest 1 mm) between the inside of
the bottom jaw and inside of the top jaw (gape height; Fig. 4). Next, measure the distance (to the
nearest 1 mm) across the width of the mouth (gape width).
Note: Gape measurements and relationships to body length may be available in published
literature in the future.
A.
B.
Fig. 4. Measuring lionfish A) gape height and B) gape width to the nearest 1 mm. Ensure that the
specimen’s mouth is open to its fullest extent.
Tissue sampling
When collecting tissues for genetic purposes, it is important not to contaminate samples.
Ensure that all dissection equipment (i.e., scissors, scalpels, forceps) are cleaned with ethanol or
DNA Zap™ (preferred) prior to taking each sample. All the tissues listed below will provide
ample DNA for genetics studies and can be placed in fixative (ethanol or DMSO buffered
solution) or frozen (Kelsch and Shields 1996). The ratio of fixative to tissue should be 50:1 for
9
proper fixation. If the tissue is intended for use in stable isotope analysis, store frozen (without
fixative) in an air-tight sample container.
Gill tissue: Using dissection scissors, remove a 1-cm2 section of gill tissue from an inside arch
including both arch and filament (Fig. 5A).
Fin clip: Remove a 1-cm2 section of pectoral fin, including membrane using dissection scissors
or a scalpel (Fig. 5B).
Muscle tissue: Using dissection scissors or scalpel, dissect a 1-cm3 section of white muscle tissue
(including skin) from the shoulder of the fish (behind the head and above the lateral line; Fig.
5C).
Fig. 5. Sample collection of three tissue types for genetic and stable isotope research: A) gill, B)
pectoral fin, and C) muscle tissue. Sample size for gill and fin tissue is 1-cm2 section; muscle
tissue sample is 1 cm3.
Internal measurements and sampling
Opening the gut cavity
Using dissection scissors, make a shallow incision from urogenital opening towards the
base of the pelvic fins (pelvic girdle) along the ventral surface of the fish (Fig. 6A). Take care to
cut shallowly; lifting up with scissors will extend the tissue away from organs helping to avoid
damage to the internal organs. As you approach the pelvic fins, a deeper cut will be required to
sever the pelvic girdle. A pair of utility scissors may be needed to sever the pelvic girdle in larger
specimens. Continue cutting anteriorly beyond the pelvic girdle to the isthmus of the gill
openings and snip through the thin flesh of the isthmus (Fig. 6B). Next, place the fish on its side
and cut along the rear edge of the gill arch toward the dorsal fin (Fig. 6C). Finally, lift the lateral
musculature to expose the gut cavity and internal organs, cutting through any minor connecting
tissue as needed (Fig. 6D).
10
Fig. 6. Opening the lionfish’s gut cavity. A) A shallow incision from urogenital opening towards
the base of the pelvic fins (pelvic girdle). B) As you approach the pelvic fins, a deeper cut will be
required to sever the pelvic girdle. C) Cutting along the rear edge of the gill arch toward the
dorsal fin. D) Lifting the flank to expose the gut cavity and internal organs, cutting through any
minor connecting tissue as needed.
11
Gender Identification and Reproductive Staging
Using a dull dissecting probe or finger, move away stomach, liver and fat to expose the
swim bladder which lies along the dorsal wall of the body cavity (Fig. 2B). Identify gonads lying
along the ventral side of the swim bladder (Fig. 2C). Avoid confusion with swim bladder muscle
tissue which lies along the dorsal edge of the swim bladder (Fig. 2D). Gonads of smaller fish
may be identified by looking for the presence of a thin blood vessel leading into and out of the
gonad. Use a magnifying glass or dissecting microscope to locate lionfish gonads in small
individuals as needed.
Using information on the size of the lionfish specimen, and the size and appearance of its
gonads greatly aids in gender and reproductive stage identification. Immature gonads from both
sexes are threadlike. As a result, testes are largely indistinguishable from immature ovaries and
may be misidentified without histology in specimens smaller than 180 mm TL (Table 2A). For
specimens greater than 180 mm TL, immature testes, appearance and size of the gonads can be
used to determine reproductive status (Table 2B).
Gonads may be removed, weighed and stored in 10% neutral buffered formalin solution for
histological examination. For long term storage, gonads that have been fixed in formalin may be
transferred to 95% ethanol. Gonads that have been previously frozen may not provide accurate
histological information because of cellular damage caused during freezing.
12
Table 2. Sex determination and gonad staging key for lionfish.
Fish length
(TL)
Appearance
Gender
Stage description
Stage
Image
Gonads are oval
masses, cream-pink
in colour, with ratio
of length:width less
than 2.
Female
Additional information required
to determine reproductive stage.
Immature
(virgin) or
Early
Developing
See Immature or Early
Developing
Gonads elongated
with ratio of length:
width greater than
2.
Histology
required to
distinguish
between male
and female.
Clear, threadlike structures 1-3
mm in diameter and 5-10 mm in
length. Immature ovaries are
largely indistinguishable from
immature testes.
Immature
(virgin)
Testes appear threadlike; 1-3
mm in diameter and 5-10 mm in
length.
Immature
(virgin)
Testes appear as cream-colored
with well-defined edges. Testes
are typically not larger than 10
mm in diameter and 20 mm in
length in the largest of
specimens.
Spawning
capable
Ovary is cream colored and
round with no edges. Width
may vary from 5 - 15 mm. Eggs
not visible macroscopically.
Ovary is more firm than during
the Developing stage.
Early
developing
Ovary cream colored with some
pinkish portions. Eggs visible as
small white spheres. Size may
vary from a width of 15 mm to
30+ mm. No gelatinous mucus
visible around periphery.
Developing
<180 mm
Gonads elongated
with ratio of
length:width greater
than 2.
Male
>180 mm
Gonads are oval
masses, cream-pink
in colour, with ratio
of length:width less
than 2.
Female
Ovary large with clear
gelatinous mucus containing
visible eggs peripheral to
central stroma. Size may vary
from a width of 15 mm to 30+
mm.
Spawning
capable
Large number of clear eggs
encompassed in gelatinous
mucus visible along periphery
of the ovary. Note: ovary wall
is removed in picture.
Actively
spawning
13
Interstitial fat deposits
Interstitial fat deposits appear as white, waxy bodies and masses around the stomach and
intestines (see Fig. 2 for orientation). Using dissecting scissors and forceps, carefully cut fat
away from the intestines and remove any other large fat deposits from the gut cavity (Fig. 7A).
Partially fill a 50-ml graduated cylinder with freshwater and note the volume at the meniscus.
Completely submerge the extracted fat deposits in the graduated cylinder and record the final
volume of fat and water. The difference between final and initial volume is the volume of the
specimen’s interstitial fat; record this value to the nearest 1ml (Fig. 7B).
Note: In fish that have warmed to room temperature, fat may begin to liquefy and become
difficult to accurately handle or remove. It is recommended that fish be refrigerated or stored on
ice prior to dissection.
Fig. 7. A) Removal of interstitial fat deposits from intestines and B) measurement of interstitial
fat volume.
Stomach contents
Before opening the stomach, ensure that prey items or parts of prey are not protruding from
the esophagus by looking into the mouth of the lionfish. Locate the stomach (Fig. 2). Using
dissecting scissors sever the esophagus where it terminates (Fig. 8A) taking care not to cut any
stomach items that may be protruding into the esophagus. Holding the esophagus closed with
fingers or forceps, cut the stomach free from other minor tissue membranes and intestines.
Remove the stomach from the body cavity and place it on a clean surface.
Insert scissors into the esophageal opening of the stomach and make a careful shallow cut
along the length of the stomach wall to the intestine opening (Fig. 8B and C). Invert the stomach
and carefully remove all prey items and place them onto clean surface for sorting and
identification. Identify each prey item to the lowest taxonomic level possible and measure the
total length of each individual prey item where possible (Fig. 8D). Use visual references guides
14
as needed to assist with visual identification of fish prey (e.g., Humann and DeLoach, 2002;
ReefNet Inc. 2007). For prey items that are too digested to discern body shape (and thus length),
length estimation is not possible. Score the digestion level of each prey item using the
description is Table 3. For all prey items, use a 10-ml graduated cylinder, measure the volume of
each prey by water displacement (same method described for measuring the volume of
interstitial fat). Preserve prey items either frozen or fixed in 95% ethanol.
Fig. 8. Stomach dissection and contents analysis. A) Using dissecting scissors sever the
esophagus where it terminates. B) Insert scissors into the esophageal opening of the stomach.
C) Make a shallow cut along the length of the stomach wall to the intestine opening.
D) Invert the stomach and remove all prey items; placing prey onto a clean surface
for sorting and identification.
15
Table 3. Five-point scale for scoring the digestion level of individual prey items removed from
the lionfish stomach.
Score
1
2
3
4
5
Description
No degradation or digestion; prey item appears freshly
consumed
Minor degradation to fin rays and scale pigmentation
Substantial degradation to external structures such as fin
rays and scale pigmentation
Major degradation: parts of body flesh missing, no
pigmentation, length measurement not possible
Prey item fully digested and reduced to mush-like
condition
Identification
resolution
Species
Species
Family
Vertebrate/invertebrate
Visual identification not
possible
Otolith removal
Otoliths, or ear stones, are small bone-like structures in the otic bulba of the cranial cavity
of fish. As a fish grows, deposits are made on otoliths in the form of rings. These rings can be
used for determining fish age and growth rates (Fig. 9A through C). Otoliths are composed of
calcium carbonate and are brittle. Care must be taken during removal to avoid damaging them.
There are three otoliths on each side of the otic bulba in the cranial cavity (grouped into pairs).
Sagittal otoliths are the largest and their dissection is described below and in Fig. 10.
Fig. 9. Ontogenetic change in sagittal otolith shape, taken from lionfish of A) 250 mm, B) 325
mm and C) 357 mm total length. Photo credit: Jennifer Potts.
16
There are several ways to perform otolith removal. Here we provide instructions for
otolith removal using a technique that involves removal of the head for easy access to the
sagittae. In some cases where trauma to the head of the carcass is an issue, other methods may
be more appropriate (e.g., trophy fish caught during tournaments). Detailed instructions for
alternate techniques of sagittal otolith removal can be found in Secor et al (1991).
Remove the specimen’s head by cutting vertically from between the head spines and the
first dorsal spine to the isthmus of the gill opening using a large, sharp knife. Turning the head
nose down following severance of the spinal cord (Fig. 10A) reduces difficulty in cutting
through cartilage. Holding the head dorsal-side down, grasp gill filaments and arches and sever
their attachment to the roof of the oral cavity (Fig. 10B). Fold the gill arches forward to reveal
the cranial cavity, which can be easily located by running two fingers along both sides of the
spinal cord anteriorly until a slight bulge is felt (the cranial cavity; Fig. 10C ). Place two fingers
on the cranial cavity and make a cut on the posterior side of the cavity down through the spine
(Fig. 10D). This incision allows access to the cranial cavity which contains the sagittal otoliths.
Proper position of this incision is critical. Cutting too far forward into the cranial cavity will
dislodge the otoliths and make them difficult to recover and cutting too shallow will prevent
access to the cavity (Fig. 10E). Using a pair of fine-tipped forceps, reach into the cranial cavity at
an oblique angle (~30º) to the dorsal plane, holding the forceps perpendicular to the dorsal plane.
Remove the otoliths by reaching into the cranial cavity. Otoliths are free-floating inside the
cranial cavity and resistance felt while searching inside the cranial cavity indicates that the
structure held is not an otolith (Fig. 10F). Once removed, clean and dry the otoliths and place in
a storage container.
17
Fig. 10. Sagittal otolith removal. A) Severe the spinal cord between the head and the
first dorsal fin ray. B) Grasp the gill filaments and arches and severe their attachment to
the roof of the oral cavity. C) Locate the cranial cavity by running two fingers along both
sides of the spinal cord anteriorly until a slight bulge is felt. D) Make a cut on the posterior
side of the cavity down through the spine. This incision allows access to the otic bulba,
which contains the sagittal otoliths. E) Brain matter (white, soft tissue) will often be
exposed by opening the cranial cavity. F) Reach gently into the cranial cavity to remove
the free-floating otoliths.
18
Acknowledgements
This work provides research findings supported by the NOAA Aquatic Invasive Species
Program, NOAA National Centers for Coastal Ocean Science, Reef Environmental and
Education Foundation, Natural Sciences and Engineering Research Council of Canada, and
Simon Fraser University. We thank N. Brown-Peterson, M. Albins, I. Côté, C. Buckel, J. Potts,
P. Marraro, M. Fonseca, and K. Riley for their helpful review of this manuscript.
19
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http://www.avma.org/issues/animal_welfare/euthanasia.pdf. Accessed January 2012.
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sting-induced paralysis. AACL International Journal of Bioflux Society 5: 1-3.
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20
Appendix I: Example template - Lionfish dissection data sheet
Lionfish ID
Date
Region
Site
TL
SL
(mm) (mm)
Weight
(g)
Sex
Gonad
stage
Fat
(ml)
Otoliths
Finclip/gill
/muscle
Dissector Name
21
Appendix II: Example template - Lionfish stomach contents data sheet
Lionfish ID
Stomach item ID
Size
(mm)
Volume
(ml)
Weight
(g)
Digestion level
(1-5)
Preserved?
22
Appendix III: Dissection Equipment List
Dissection Equipment List
*quantity of each item will vary with number of specimens
Puncture-resistant gloves
Euthanasia
Per 10 L seawater:
5 ml Clove oil
45 ml Isopropyl or ethyl alcohol
OR
4 L ice
Dissection
95% Ethanol
Formalin
Lionfish specimen data sheet
Lionfish stomach contents data sheet
Large dissection board
Large sharp kitchen knife
Fine-tipped forceps (2 pair)
Dissecting scissors (1 pair)
Paper towels
Volumetric flasks (10 ml and 50 ml)
Pipette
Sharpie permanent pen
Specimen sample labels (Underwater paper cut into 2” x 2” sections)
Screw-capped glass sample vials (~5 ml volume)
Whatman filter paper
Small plastic sample bags (~10 ml volume)
Vials or envelopes for otoliths
23
Appendix IV: Lionfish Tissue Repository
What is the Lionfish Tissue
Repository?
The Lionfish Tissue Repository (LTR) is a
multi-national collaborative program
intended to maintain and provide tissue
samples for research into the ecological and
evolutionary processes driving the ongoing
invasion of lionfish (Pterois spp.) in the
Caribbean and western Atlantic.
Who contributes to repository?
Scientists, managers and fishers throughout the western Atlantic have contributed tissue samples
to the growing database. Large collections of samples from the eastern United States, Bahamas
and Cayman Islands have been contributed by organizations such as NOAA, Reef Environmental
Education Foundation (REEF), Oregon State University, George Washington University, and the
Cayman Islands Department of the Environment.
Who manages the repository?
The repository is jointly managed by NOAA and Reef Environmental Education Foundation
(REEF). There are currently well over 3,000 genetic tissue samples spanning nearly a decade
collected from lionfish throughout the invaded Caribbean and eastern coast of the US. As
samples continue to accumulate, we expect the tissue repository will yield a uniquely detailed
genetic and ecological history of a major invasion.
Who is using the repository?
The intent of the LTR program is to carry out collaborative research that will help scientists and
natural resource managers understand the invasion (and invasion ecology/evolution in general),
identify mechanisms for mitigating the invasion impact, and prevent future marine invasions. We
anticipate that the LTR will be most useful for the study of lionfish genetics or foraging ecology
(e.g. stable isotope ecology). There may, however, be additional applications not yet considered.
By maintaining the LTR, we preserve the opportunity to address future research questions in an
opportunistic manner.
For additional questions about on-going research, contact the project principal investigator Dr.
James Morris at James.Morris@noaa.gov or visit
http://www.ccfhr.noaa.gov/stressors/lionfish.aspx.
24
United States Department of Commerce
John E. Bryson
Secretary
National Oceanic and Atmospheric Administration
Jane Lubchenco
Undersecretary of Commerce for Oceans and Atmosphere
and NOAA Administrator
National Ocean Service
David Kennedy
Assistant Administrator for Ocean Services and Coastal Zone Management